Chromatin Immunoprecipitation followed by qPCR and sequencing

In the field of gene expression, we are often concerned with the genomic location of a given factor and the identity of sequences that it associates with. Knowing the genomic binding sites of a protein of interest, or the location of specific histone modifications, often gives us valuable information about global patterns of gene regulation.

In vivo crosslinking with formaldehyde and chromatin immunoprecipitation (ChIP) has emerged as a widely used and important technique in the analysis of protein localization in a physiological setting. This technique relies on the ability of formaldehyde to crosslink proteins with other proteins and proteins with DNA. After chromatin is immunoprecipitated with antibodies directed against the target protein of choice, cross-links can be reversed with heat treatment. DNA fragments isolated in this way can then be detected by PCR amplification, Southern blotting, oligonucleotide microarrays or deep sequencing. The versatility of the ChIP technique is illustrated by the fact that it has been used successfully to study proteins recruited directly to the DNA, as well as proteins that are recruited to promoters indirectly through intermediary factors. Furthermore, ChIP permits the study of histone modifications, such as acetylation or methylation, at specific sequences, especially promoters. The modification of histones at promoters is a central issue in the gene expression field. Furthermore, covalent modifications of other chromatin-bound factors, such as Pol II itself, can be also analyzed with this technique. In this set of experiments, we will use ChIP assays to define the characteristic pattern of Pol II distribution and histone occupancy at active and inactive genes.

We will perform both ChIP-seq and ChIP qPCR (used layout). Each bay (group of 4 people) should pick one group of 2 to carry out Pol II IPs for Pol II ChIP-seq, and one group of 2 to carry out Pol II and histone IPs for analysis by qPCR with an Absolute Quantification method.

Outline:

A. Preparation of crosslinked protein extract from Drosophila S2 cells

B. Sonication of Chrmatin Immunoprecipitation (ChIP) material

Sonication of ChIP material using Bioruptor and checking fragment sizes

C. ChIP Analysis by qPCR using tiled primer sets

Pre-clear ChIP material
Immunoprecipitation with specific antibodies and controls
Washes and elution of immunoprecipitated material
DNA precipitation and preparation for qPCR
Master Mixes, Input Genomic DNA Standard Curve
Samples and Layout

D. ChIP-seq library preparation

Pre-clear ChIP material
Immunoprecipitation with specific antibodies
Washes and elution of immunoprecipitated material
DNA precipitation and library preparation

E. Solutions and Reagents

A. Preparation of crosslinked protein extract from Drosophila S2 cells

We will perform ChIP on Drosophila S2 cells grown in Schneider’s media + 10% FBS.
Example volume below are for one T175 cm^2 flask, with ~30 mL total volume.

S2 cells are plated at a low cell density (1 x 10^6 cells/mL) in 30 mL of media in a T175 flask.
– Let cells grow until they reach a density anywhere between ~ 4-8 x 10^6 cells/mL.

This represents 2 to 3 doubling events, and should take ~ 3 days.
When cells appear to be at appropriate confluency (after ~ 3 days), take an aliquot to determine the exact number of cells you will be crosslinking.
– Tap the flasks against your hand and squirt the cells from the surface using a 10 mL pipette if necessary to suspend the cells in media.
– Taking care to swirl to remove to remove a homogeneous suspension of cells, remove a small aliquot (~ 500 uL) of cells from each of your flasks for determining cell count.
– Mix 10 uL of cells with 10 uL of trypan blue and count live cells using BioRad TC20.
The cell density, times 25 mL volume used below, will determine the volume of sonication buffer in which each sample is resuspended later on.
Remove 25 mL of cells from T175 flask and mix with 50 mL room temperature 1X PBS in a 250 mL Erlenmeyer flask.
– Swirl briefly.
This dilutes the serum and allows for good mixing with formaldehyde so that crosslinking is efficient.
Crosslink cells for 5 min with a final concentration of formaldehyde of 1 %
Cells in 250 mL Erlenmeyer flask are at room temperature on the bench top.
– Make up 11% Formaldehyde working solution right before use.
Mix 6 mL H2O + 1 mL 10X PBS + 3 mL 37% formaldehyde
Note: remember that the formaldehyde stock is 37%, not 100%For 75 mL of cell volume in this example, add 7.5 mL 11% Formaldehyde solution.
This gives you a final formaldehyde concentration of 1%
Quench Cross-linking
Add 4.5 mL Glycine (2.5M stock solution) to ~ 90 mL of solution in the Erlenmeyer flask.
This represents a 1:20 dilution, or a final concentration of 125 mM to quench crosslinking.
– Swirl sample with glycine to mix well.
Isolating cross-linked material
– Cell suspension is transferred to 2 x 50 mL screw cap conical tubes, and placed on ice to cool for 2-3 min.
– Pellet cells at 1000 x g for 3 min at 4C in Beckman Allegra centrifuge or similar.
– Carefully dump off supernatant.
– Spin again to remove all media.
Rinse the cells and prepare the pellet for sonication
– Resuspend/rinse the pellet with 10 mL 1x PBS and transfer to a 15 mL conical tube.
This will remove residual media and serum that might interfere with downstream application.
– Spin again at 1000 x g and carefully aspirate supernatant. Place the cell pellet on ice.
Lyse the cells in sonication buffer
Resuspend each pellet in 1 mL cold sonication buffer per 1 x 10^8 cells
– Calculate the total number of cells in your sample, and adjust the sonication buffer volume accordingly
– Keeping a constant concentration of ChIP material across all samples greatly increases the reproducibility in IPs.
The sonication buffer should be ~ 1000-1500 uL/ per a standard 25 mL cell culture.Sonication buffer
final concentration for 10 mL
0.5% – 500 uL 10% SDS
20 mM – 200 uL 1M Tris, pH 8.0
2 mM – 40 uL 0.5M EDTA
0.5 mM – 25 uL 0.2M EGTA
0.5 mM – 50 uL 100mM PMSF
9.15 mL H2O
1 Complete Protease Inhibitor tablet (Roche, Complete Mini)

The sonication buffer should be made fresh so that SDS doesn’t precipitate or PMSF goes off.
– After gentle, but thorough resuspension of the cell pellet, incubate the cells on ice for 10 min to allow cell lysis.
– Aliquot ChIP material and flash freeze. Store at -80C until use.

B. Sonication of Chromatin Immunoprecipitation (ChIP) material

Sonicate cross linked material using a Bioruptor Plus.
– Add 500 mg of sonication beads from Diagenode to 15 mL sonication tube (also from Diagenode).
These beads are light and difficult to weigh, so be careful on the balance and transferring into the sonication tube.
– Add 3 volume (~ 600 uL) of 1X PBS
– Vortex well and centrifuge 500xg for 2 min.
– Remove all 1xPBS. This can be tricky. Please ask your TA if you need help.
– Add 1 mL of ChIP material to the washed sonication beads.
– Do 25-30 pulses of sonication.
Each pulse of sonication is for 30″ with 1 min rests between each pulseNote: For these cells, this amount of sonication should give you fragment sizes of between 200-500 bp. If your fragments are bigger, you should sonicate more… this part is going to be different for different cell types and sonication systems and needs to be determined empirically.
Remove the cellular debris
Spin sample quickly in benchtop centrifuge to collect material splasjed on the sides of the tubes (e.g. spin at 500 x g for 1 min).
Transfer sonicated material to a 1.5 mL Eppendorf tube and spin at maximum speed for 10 min at 4C to remove cell debris.Transfer supernatant (carefully) to a fresh 1.5 mL Eppendorf tube. This contains solubilized crosslinked material for immunoprecipitations.

Take an aliquot of 25 uL to check the fragment size.

The remainder should be flash frozen and stored away at -80C.
Note: We will do the IPs from these samples the following day, but they can be stored for several months before use.
Reverse crosslinks to evaluate the DNA fragment size
Make the Elution/cross-link reversal solution which is 1% SDS and 0.1 M NaHCO3 (10 mL = 8 mL H2O + 1 mL 10% SDS and 1 mL 1 M NaHCO3)To each 25 uL aliquot of ChIP material, add:
500 uL Elution solution
+ 20 uL 5M NaCl
Heat at 65C in a water bath for > 3.5 hours to reverse crosslinks.
Deproteinate to evaluate the DNA fragment size
After > 3.5 hours at 65C, remove the sample from the water bath and spin for 30 s in a microfuge.
Do this BEFORE you open the tube, to avoid all the condensed liquid on the tube lid from flying away!Add to each sample:
15 uL 1 M Tris pH 7.8
2 uL Glycoblue
2 uL Proteinase K
Let it incubate for at least 30 more min at 65C
Note: This digests up protein and yields a much nicer, cleaner interface.
Phenol/chloroform extraction
Take the samples from the 65C bath and spin for 30 s in a microfuge.
Add 500 uL phenol/chroroform to each sample.
– Phenol/chloroform is kept in the fume hood, and added to the tubes in the fume hood.
Vortex each sample well.Spin for 10 min in a room temperature centrifuge at maximum speed.

Carefully remove the tubes from the centrifuge.
– Remove 450 uL of upper, aqueous layer to a fresh tube.
– Dispose of the bottom layer appropriately in the fume hood.
Precipitate the DNA
Precipitate the DNA by adding 1 mL room temperature EtOH to your 450 uL DNA in aqueous phase.
Be careful not to cool the samples, or SDS will precipitate!Let the samples sit for > 30 min.

Spin the samples for 20-30 min at maximum speed.
Wash the pellets carefully with 175 uL 70% EtOH, spin again for 5 min.
Let the pellets air dry.
Run the DNA on a agarose gel
Resuspend each DNA pellet in 20 uL H2O plus:
3-5 RNase cocktail and 1-2 uL RNaseH (Ambion).Let it sit at RT for at least 30 min to degrade any RNA.

Add the sample loading buffer (blue stuff, obtained from the TA).

Run 20 uL on 1% agarose gel to check the average fragment size of DNA, use a marker that gives good resolution between 100 and 1000 bp.

C. ChIP Analysis by qPCR using tiled primer sets

In this section, you will analyze your ChIP-enriched DNA by performing quantitative real-time PCR (qPCR), with products visualized by the intercalating dye SYBR-Green. The critical aspect of this procedure is the ability to compare relative enrichments (i.e. amount of amplification from IP and control samples) for a given test sequence with those from a negative control sequence and the use of genomic input DNA to calculate an absolute Percentage of Input recovered in each immunoprecipitated sample.

Each group will be provided with primer sets that will amplify the relevant sequences along heat shock genes and a control gene; these primers are generally 18-24 bp in length with a Tm of 60+/-5 degrees Celsius.
Each new primer set should be tested before use by amplification an “Input” DNA sample.
– We will use previously validated primer sets for the course.

Setting up an immunoprecipitation
INPUT: You need to take an input sample every time you do an IP so that you can compare the yield in the qPCR to the total amount of genomic DNA present in that particular ChIP preparation.

This should usually be done in duplicate, unless this makes just too many samples to handle.
Take an aliquot of the cell sample that will equal ~ 20% of the cells used/IP
For example, if you are using 25 uL ChIP material per IP, take two aliquots of 5 uL (or 20% input matertial).

Pre-clear ChIP material
– Pre-clear each ChIP sample with Protein-A agarose equilibrated in IP buffer. This gets rid of sticky junk that will adhere to the beads even in the absence of antibodies.
You will use 25 uL ChIP material per IP (= 2.5 x 10^6 cells/IP)
– We will dilute this into 1 mL total volume with IP buffer.
To this, we will add 30 uL of a 50% slurry of Agarose beads per sample.For multiple samples from the same material, you can pre-clear in bulk in a 15 mL tube.
Always prepare one more sample than you need!

e.g. for 6 IPs, pre-clear is made up for 7 samples:
25 uL ChIP material x 7 = 175 uL
30 uL Protein-A agarose slurry x 7 = 210 uL
945 uL IP buffer x 7 = 6.615 mL
total = 7 mL

– Leave them for 1-2 hours at 4C on the rotating wheel.
After you have set up the pre-clear, place the remaining ChIP material at -80C.

– Spin the samples for 1 min at 1000 xg in the Allegra centrifuge.
Immunoprecipitation with specific antibodies and controls
Aliquot 1 mL supernatant from pre-clear into each of 6 Eppendorf tubes (1mL/tube).
To each tube, add an additional 250 uL IP buffer.
NOTE: It is important to dilute ChIP material for the IP, because the SDS can interfere with antibody interactions.Add immunoprecipitating antibody in appropriate amount.
2 Rpb3 IPs (6 uL antibody / IP)
2 Histone H3 IPs (15 uL antibody / IP)
1 Histone H3K4me3 IPs (15 uL antibody / IP)
no antibody control

Place IP material at 4C on rotating wheel overnight.

The next morning, add Protein-A agarose (50% slurry), equilibrate in IP buffer.
200 uL per sample, to ensure about 100 uL of bead volume
– After adding slurry, let the beads settle for a minute to ensure that you have sufficient, and equal, bead volume in each tube.

Let the material incubate at 4C on rotating wheel for 1-2 hours.
Washes and elution of immunoprecipitation material
Ideally, you would spin the samples in pre-chilled (4C) microfuge at 1000 xg (3500 rpm) for 1 min to pellet the beads.
But, room temperature centrifugation is fine if you don’t have access to temperature-controlled centrifuges. Just work quickly!
– All spins are 1000 xg (350 rpm) for 1 min
– Remove supernatant to trash, and begin washes of beadsWashes
Each wash is 1 mL volume of ice cold wash solution, and is left for 3 min on rotating mixer at room temperature, then is spun for 1 min at 1000 xg.
Do 1 Low salt wash, 3 High salt washes and 1 LiCl wash.

TE equilibration
Rinse beads 2 times with 1mL TE, mix and spin as above.

Elution
Elute DNA with 250 uL freshly prepared Elution solution (1% SDS and 0.1 M NaHCO3)
(20 mL = 16 mL H2O + 2 mL 10% SDS and 2 mL 1 M NaHCO3)

Let this elution buffer + beads sit on rotating mixer for 15 min,
Spin as above and remove supernatant to fresh tube as your eluted material.

Second elution
Add another 250 uL Elution solution and repeat elution for 15 min incubation, spin
Combine the second eluates with the first for total volume = 500 uL
DNA precipitation and preparation for qPCR
Reverse cross-links and isolate DNA
Reverse cross-links on input material too
Add 500 uL Elution buffer and 20 uL 5M NaCl to the input material you froze away the day before.Add 20 uL 5M NaCl to each eluted ChIP sample
Heat all samples at 65C for 4 hours to reverse crosslinks

Deproteinate. After > 3.5 hours at 65C, add 15 uL 1M Tris pH 7.8, 2uL Glycoblue + 2 uL Proteinase K to each sample, let incubate for at least 30 min (this digests up protein and yields a much nicer, cleaner interface).

Phenol/Chloroform extraction of each sample.

Precipitate DNA samples with 1 mL room temperature EtOH or equal volume isopropanol
Be careful not to cool sample, or SDS will precipitate!
– Let samples sit for 30 min to overnight.

Spin for 20-30 min at maximum speed.
Wash with 70% EtOH, let pellet air dry.

Resuspend DNA.

If you will be doing qPCR, resuspend each pellet in 100 uL H2O
– You will perform PCR reactions with 4 uL of each ChIP sample and serial dilutions of the input DNA.
Master Mixes, Input Genomic DNA Standards
For a 384-well plate the final volume in each well is 20 uL.
4 uL of the immunoprecipitated DNA or genomic input DNA standards are pipetted into each well.
The SYBR Green master mix + primers is 16 uL per well and this Master Mix is made up in bulk.Reaction Master Mix for each primer pair (e.g. for 50 reactions):
1.33 uL Forward primer (5 uM stock) x 50 = 66.5 uL
1.33 uL Reverse primer (5 uM stock) x 50 = 66.5 uL
10 uL iTaq Universal SYBR Green mix x 50 = 500 uL
3.4 uL H2O x 50 = 170 uL
~ 16 uL per sample

Input DNA serial dilutions:
Resuspend input in appropriate volume for serial dilutions from 10% to 0.01%
e.g. resuspend 10% input, which is 7.5 x 10^5 worth of cell material, in the same volume of H2O as immunoprecipitated samples (i.e. 200 uL). This is now your “10% input” material.

Make serial dilutions of 10% input material (1:100, 1:1000, 1:10,000) by serially transferring 20 uL into 180 uL H2O, for 200 uL total of each dilution.

Make sure you amplify 10%, 1%, 0.1% and 0.01% inputs for each sample with each primer set used to generate standard curves. Ideally, you will have 2 separate inputs or at the very least triplicate PCRs of each input and will really be sure of this standard curve.
Samples and Layout
Samples are typically diluted to fall somewhere in the middle of the standard curve. This should be determined empirically depending on a number of variables, such as antibody quality, protein-chromatin interaction, etc.See the layout of PCR plate provided by your TA for the details of which samples goes where.

Make sure to use a background primer set (negative control) and a number of gene regions for qPCR reactions.

Once your layout is set, prepare the Master Mixes for each primer pair first.
Then transfer DNA samples to appropriate well of PCR plate in 4 uL per sample.
Finally, 16 uL of the appropriate master mix is aliquoted into each well, often using a repeater pipette.

Spin the plate down at 500-1000 rpm for about 1 min
(I usually leave the plate in the fridge covered with foil for 50-60 min to decrease background fluorescence from exposure of Sybrgreen to light during plate loading. If I do this, I spin the plate before putting it in the machine to also minimize condensation)

Run standard amplification (Taqman) protocol
50C for 2 min
95C for 10 min
40 cycles of:
94C 15 sec, then 60C for 1 min, which is annealing and extension time

Don’t forget to add dissociation curve so that you can assay the DNA amplified.

D. ChIP-seq library preparation

Pre-clear ChIP material
– Pre-clear each ChIP sample with Protein-A agarose equilibrated in IP buffer.
This gets rid of sticky junk that will adhere to the beads even in the absence of antibody.For ChIP-seq, you will perform 6 IPs, using 75 uL ChIP material each (= 7.5 x 10^6 cells/IP)
– We will pre-clear this in 1 mL total volume, by diluting with IP buffer.
– We will add 100 uL of a 50% slurry of Protein-Agarose beads per sample.

For multiple samples from the same material, you can pre-clear in bulk in a 15 mL tube.
Always prepare 1 more sample than you need!

e.g. for 6 IPs, pre-clear is made up for 7 samples:
75 uL ChIP material – x7 = 525 uL
100 uL Protein-A agarose slurry – x7 = 700 uL
825 uL IP buffer – x7 = 5.775 mL
= 7 mL total

– Leave it for 1-2 hours at 4C on the rotating wheel
After you have set up pre-clear, place the remaining ChIP material at -80C

– Spin the samples for 1 min at 100 xg in the Allegra centrifuge.
Immunoprecipitation with specific antibodies
Aliquot 1 mL supernatant from pre-clear into each of 6 Eppendorf tubes (1 mL/tube).
To each tube, add an additional 250 uL IP buffer.
Note: It is important to dilute the ChIP material for the IP, because the SDS can interfere with antibody interactions.Add immunoprecipitating antibody in appropriate amount.
15 uL Rpb3 antibody per IP

Place IP material at 4C on rotating wheel overnight

The next morning, add Protein-A agarose (50% slurry), equilibrated in IP buffer.
200 uL per sample, to ensure about 100 uL of bead volume.
– After adding slurry, let the beads settle for a minute to ensure that you have sufficient, and equal, bead volume in each tube.

Let the material incubate at 4C on rotating wheel for 1-2 hours.
Washes and elution of immunoprecipitation material
Ideally, you would spin the samples in a pre-chilled (4C) microfuge at 1000 x g (3500 rpm) for 1 min to pellet the beads.
But, room temperature centrifugation is fine if you don’t have access to temperature-controlled centrifuges. Just work quickly!
– All spins are 1000 x g (3500 rpm) for 1 min
– Remove supernatant to trash, and begin washes of beadsWashes
Each wash is 1 mL volume of ice cold solution, and is left for 3 min on a rotating mixer at RT, then is spun for 1 min at 1000 x g.

Do 1 Low salt wash, 3 High salt washes and 1 LiCl wash.

TE equilibration
Rinse the beads 2 times with 1 mL TE, mix and spin as above.

Elution
Elute the DNA with 250 uL freshly prepared Elution solution (1% SDS and 0.1 M NaHCO3)
(20 mL = 16 mL H2O + 2 mL 10% SDS and 2 mL 1 M NaHCO3)

Let this elution buffer + beads sit on a rotating mixer for 15 min,
Spin as above and remove the supernatant to a fresh tube as your eluted material.

Second elution
Add another 250 uL Elution solution and repeat the elution for 15 min incubation, spin
Combine the second elutes with the first for a total volume = 500 uL
DNA precipitation and library preparationReverse cross-links and isolate the DNA:
Reverse cross-links on input material too
Add 500 uL Elution buffer and 20 uL 5M NaCl to the input material you froze way the day before.

Add 20 uL 5M NaCl to each eluted ChIP sample
Heat all samples at 65C for 4 hours to reverse the crosslinks

Deproteinate. After > 3.5 h at 65C, add 15 uL 1 M Tris pH 7.8, 2 uL Glycoblue + 2 uL Proteinase K to each sample, let in incubate for at least 30 min (this digests up the protein and yields a much nicer, cleaner interface).

Phenol/chloroform extractions of each sample

Precipitate DNA sample with 1 mL room temperature EtOH or equal volume isopropanol.
Be careful not to cool the samples, or SDS will precipitate!
– Let the samples sit for 30 min to overnight.

Spin for 20-30 min at maximum speed.
Wash with 70% EtOH, let pellets air dry.

Resuspend DNA
– For making ChIP-seq libraries, resuspend each pellet in 20 uL H2O
– When you combine all 6 IPs below, this will give you 120 uL final volume

Libraray preparation
We will perform this with a kit by Rubicon Genomics.
We will follow the protocol provided with this kit, followed by a gel purification / size selection step.

a) Sample preparation / cleanup:
– Pool all 6 samples into one 1.5 Eppendorf tube and add 5 volumes (600 uL) PB buffer
– Transfer the sample to MinElute Spin column and spin for 30 sec at 16000xg
– Add flow through again to the binding column and spin for 30 sec at 16000xg
– Discard flow through, add 700 uL of PE buffer and spin for 30 sec at 16000xg
– Discard flow through and spin for 1 min at 16000xg
– Discard the collection tube and transfer the binding column to the 1.5 mL tube
– Add 10 uL or RNAse-free H2O to the column membrane and wait 1 min.

– Spin 16000xg for 1 min
– Add additional 6 uL of RNAse-free H2O to the column membrane, wait 1 min and spin at 16000xg for an additional min.

b) Quantification of eluted IPed DNA with Qubit

The TA will prepare and provide Qubit working solution in a 0.5 mL tube to each group, so that all samples are quantified simultaneously. The TA will also prepare the standards. Each group should do steps highlighted in bold. The protocol to prepare Qubit working solution is described below for reference.

– At this step, each sample requires 199 uL of working solutions, so prepare enough working solution in batch for 2 standards plus the number of samples needed.
– Label 0.5 mL tubes, two for the standards and the remaining for your samples.
– Prepare enough working solution for all samples by diluting Qubit dsDNA HS reagent 1:200 in Qubit dsDNA HS buffer.
– Add 190 uL of Qubit working solution to each of the 0.5 mL tubes labeled standards.
– Add 10 uL of each Qubit standard and vortex for 2-3 sec.
– Add 199 uL of Qubit working solution to each of the 0.5 mL tube samples.
– Add 1 uL of each sample to the 0.5 mL tube labeled for your sample and vortex for 2-3 sec.
– Allow all tubes to incubate for 1-2 min and proceed to measure the concentrations, as demonstrated by your TA.

c) Library preparation with Rubicon ThruPLEX DNA-seq kit

– For full instructions of library preparation kit see:
http://rubicongenomics.com/wp-content/uploads/2016/09/ThruPLEX-DNA-seq-Kit-Manual-QAM-108-003-updated-9_16_16.pdf (link not working)
alternative: SMARTer ThruPLEX DNA-seq Kit User Manual
1. 1. Dilute 10 ng of IPed DNA in 10 uL H20
    NOTE: The maximum volume of DNA cannot exceed 10 uL.
Template Preparation Master Mix (End-Repair and A-tailing)
1. 1. Prepare Template Preparation Master Mix as described in the table below for the desired number of reactions. Mix thoroughly with a pipette. Keep on ice until used.Component – Volume / Reaction
    Template Preparation Buffer – 2 uL
    Template Preparation Enzyme – 1 uL
  2. Assemble the Template Preparation Reaction Mixture as shown in the table below. To each 10 uL sample from step 1 above, add 3 uL of the Template Preparation Master Mix.Component – Volume / Reaction
    Sample – 10 uL
    Template Preparation Master Mix – 3 uL
    Total Volume – 13 uL
    
    – Mix thoroughly with a pipette.
    – Place tube in a thermal cycler with heated lid 100-105C.
    
    Perform the template Preparation Reaction using the conditions in the table below:
    
    Temperature – Time
    22C – 25 min
    55C – 20 min
    4C – Hold < 2 hours
Library Synthesis Reaction Mixture (Adaptor ligation)
1. 1. Prepare Library Synthesis Master Mix as described in the table below for the desired number of reactions. Mix thoroughly with a pipette. Keep on ice until used.Component – Volume / Reaction
    Library Synthesis Buffer – 1 uL
    Library Synthesis Enzyme – 1 uL
    
    – Assemble the Library Synthesis Reaction Mixture as shown in the table below.
    To each well or tube, add 2 uL of the Library Synthesis Master Mix.
    
    Component – Volume / Reaction
    Template Preparation Reaction Product – 13 uL
    Library Synthesis Master Mix – 2 uL
    Total Volume – 15 uL
    
    – Mix thoroughly with a pipette.
    – Place tube in a thermal cycler with heated lid 100-105C.
    
    Perform the Library Synthesis Reaction using the conditions in the table below:
    
    Temperature – Time
    22C – 40min
    4C – Hold < 30min
Library Amplification
1. Prepare the Library Amplification Master Mix as described in the table below for the desired number of reactions.
  Mix thoroughly with a pipette.
  Keep on ice until used.Component – Volume / Reaction
  Library Amplification Buffer – 25 uL
  Library Amplification Enzyme – 1 uL
  H2O – 4 uL
  
  – Add 30 uL of the Library Amplification Master Mix to each well or tube.
  – Add 5 uL of the appropriate Indexing Reagent to each well or tube.
  
  Component – Volume / Reaction
  Library Synthesis Reaction Product – 15 uL
  Library Amplification Master Mix – 30 uL
  Indexing Reagent – 5 uL
  Total Volume – 50 uL
  
  NOTE: The Indexing Reagents consists of amplification primers containing Illumina-compatible indexes. In order to run multiple samples in one single sequencing run each group will amplify their samples with one unique index (this will be chosen by the TA).
  
  – Return the tube(s) to the real-time PCR thermal cycler / thermal cycler with heated lid set to 100C-105C. Perform Library Amplification Reaction using the cycling conditions in the table below.
  
  Notes: The number of PCR cycles required at Stage 5 of the Library Amplification Reaction is dependent on the amount of input DNA and thermal cycler used. Use the table below as a guide for selecting the number of PCR cycles.
  
  DNA Input (ng) – Number of Cycles
  50 – 5
  20 – 6
  10 – 7
  5 – 8
  2 – 10
  1 – 11
  0.2 – 14
  0.05 – 16
  
  Optimization experiment: Performing an optimization experiment to identify the appropriate number of PCR cycles needed is recommended. Use the desired amount of input DNA and allow the library amplification reaction to reach plateau. Determine the optimal number of amplification cycles by constructing PCR curves and identifying the midpoint of the linear phase as illustrated in Figure 5. Use the optimal amplification cycle number in the actual experiment for sequencing.
  
  Yield: The amount of amplified library can range from 100 ng to 1 ug depending upon many variables including sample type, fragmentation size, and thermal cycler used. When starting with Covaris-fragmented reference DNA with an average size of 200 bp and following the recommended number of amplification cycles, the typical yield range from 300 ng to 700 ng.
  
  Note: Over amplification could result in higher rate of PCR duplicates in the library.
  
  Library Purification
  
  – Add 10 uL 6xDNA loading dye and load on two adjacent lanes of a 6% TBE gel, alongside 10 uL of DNA ladder
  – Run the gel at 200 V in 1xTBE for 25-30 min or until the bromophenol blue dye is ~ 1 cm from the bottom.
  – Stain the gel with ethidium bromide as above and excise the library, which should be clearly visible in the size ranges of 200-600 bp and separated from a linker-linker ligation product, which runs at ~75 bp
  – Cut the library out of the gel, trying not to take any of the linker-linker product
  – Transfer gel to 2 ml tube and elute in 400 uL 1xTE buffer supplemented with 50 mM NaCl for at least 2 hours
  – Transfer the entire content of the 2 mL tube into a spin column and spin at 1000xg for 2 min
  – Discard the column containing gel fragments. To the eluate at the bottom of the tube, add 1.5 uL glycoblue, 40 uL 3M NaAcetate, and 2.5 volumes (1 mL) ethanol
  – Spin immediately at 16.000xg in a refrigerated microcentrifuge, wash pellet with 70% ethanol and resuspend DNA in 100 uL H2O
  – Add 5 volumes (500 uL) PB buffer
  – Transfer sample to binding column and spin for 30 sec at 16.000xg
  – Add flow through again to binding column and spin for 30 sec at 16.000xg
  – Discard flow through, add 700 uL of PE buffer and spin for 30 sec at 16.000xg
  – Discard flow through and spin for 1 min at 16.000xg
  – Discard collection tube and transfer binding column to the 1.5 mL tube.
  – Add 10 uL of RNAse-free H2O to column membrane and wait 1 min.
  – Spin 16.000xg for 1 min.
  – Add additional 6 uL of RNase-free H2O to the column membrane, wait 1 min and spin at 16.000xg for an additional minute.
  
  Quantification of library DNA with Qubit (each sample requires 199 uL of working solution): The TA will prepare and provide Qubit working solution in a 0.5 mL tube to each group, so that all samples are quantified simultaneously. The TA will also prepare the standards. Each group should do steps highlighted in bold. The protocol to prepare Qubit working solution is described below for reference.
  
  – At this step, each sample requires 199 uL of working solution, so prepare enough working solution in the batch for 2 standards plus the number of samples needed.
  – Label 0.5 mL tubes, two for the standards and the remaining for your samples.
  – Prepare enough working solution for all samples by diluting Qubit dsDNA HS reagent 1:200 in Qubit dsDNA HS buffer.
  – Add 190 uL of Qubit working solution to each of the 0.5 mL tubes labeled standards.
  – Add 10 uL of each Qubit standard and vortex for 2-3 sec
  – Add 199 uL of Qubit working solution to each of the 0.5 mL tube samples.
  – Add 1 uL of each sample to the 0.5 mL tube labeled for your sample and vortex for 2-3 sec.
  – Allow all tubes to incubate for 1-2 minutes and proceed to measure the concentrations, as demonstrated by your TA.

E. Solutions and Reagents

37% formaldehyde stock solution (~ 12.3 M is about 37%) is diluted to 11% final concentration in PBS, using 10X PBS stock and H2O.

Glycerin (500 mL) at 2.5 M is 93.84 g Glycerin into 500 mL H2O (M.W. = 75.07).
Autoclave if you will keep it for a while.

Sonication buffer
final concentration for 10 mL
0.5% – 500 uL 10% SDS
20 mM – 200 uL 1 M Tris, pH 8.0
2 mM – 40 uL 0.5 M EDTA
0.5 mM – 25 uL 0.2 M EGTA
0.5 mM – 50 uL 100 mM PMSF
9.15 mL H2O
1 complete protease inhibitor tablet (Roche, Complete Mini)

IP buffer
final concentration for 500 mL
0.5% – 2.5 mL 100% Triton X-100
2 mM – 2.0 mL 0.5 M EDTA
20 mM – 10 mL 1 M Tris pH 8.0
150 mM – 15 mL 5 M NaCl
10% – 50 mL 100% glycerol
420.52 mL H2O

Share this: